Anal. Chem. 2001, 73, 3107-3111
Measurement of Insecticide Uptake and Effective Fraction in a Beneficial Insect Using Solid-Phase Microextraction Anne Alix,*,†,§ Damien Collot,‡ Jean-Pierre Ne´non,§ and Jean-Pierre Anger†
Laboratoire de Toxicologie Analytique, Faculte´ de Pharmacie, avenue du Professeur Le´ on Bernard, 35042 Rennes Cedex, France, CREL, De´ partement Chimie-Physique, 168, rue de Versailles, 78150 Le Chesnay, France, Laboratoire d’Ecobiologie des Insectes Parasitoı¨des, Campus de Beaulieu, avenue du ge´ ne´ ral Leclerc, 35042 Rennes Cedex, France
The determination of insecticide uptake in beneficial insects is important for quantifying the doses that are responsible for the toxicological effects and to compare them with the doses that insects may absorb in treated fields. Because of the small size of some beneficial species, the amount of insecticide absorbed may be very low. Herein, we present a method that relies on the sensitivity and specificity of SPME (solid-phase microextraction) as a sampling technique that can be used to measure very small amounts of an organophosphorus insecticide in small insects. In our study, the method was applied to quantify the internal dose and free dissolved fraction of chlorfenvinphos in beneficial parasitoids exposed through a topical application. Up to 0.5 ng of the insecticide could be quantified in these fractions, that is, 10 times less than when using solvent extraction techniques. The penetration and elimination rates of the insecticide in the insect were also determined. The method proved to be suitable to quantify internal doses in parasitoids collected in a treated field. Beneficial insects play a significant role in the control of pest populations in a number of crops1. As a consequence, their use in combination with chemical treatments has been widely practiced, because it greatly contributes to the reduction of pesticide application.1-3 The efficacy of these integrated pest management (IPM) programs clearly depends on the dose of the chemical that the beneficial species may absorb in treated fields and the toxicological effects that the absorbed dose may induce in the insect. Unfortunately, the assessment of the risk posed to beneficial insects by insecticides remains complex, mainly because the doses that these insects really absorb either in the field or in the laboratory have never been quantified.4,5 * To whom correspondence should be addressed, at: SSM INRA-DGAL, Route de St Cyr 78026, Versailles Cedex, France. Tel: (+33) (0)1 30 83 32 91. Fax: (+33) (0)1 30 83 31 49. E-mail:
[email protected]. † Laboratoire de Toxicologie Analytique, †Faculte ´ de Pharmacie. ‡ CREL. § Laboratoire d’Ecobiologie des Insectes Parasitoı ¨des. (1) Batra, S. W. T. Science 1982. 215, 134-139. (2) Brown, R. A. Pesticides and Nontarget Invertebrates; Jepson, P., Ed.; Intercept Ldt, Wimborne, U.K.; 1989, Chapter 2. (3) Hoy, M. Introduction to Insect Pest Management; 3rd ed.; Metcalf, R. L., Luckmann,W. H. Eds; John Wiley and Sons: New York, 1994, Chapter 4. 10.1021/ac0013148 CCC: $20.00 Published on Web 06/02/2001
© 2001 American Chemical Society
The quantification of insecticide uptake in insects poses many analytical problems due to body size (less than one millimeter for some species) and the complexity of the insect matrix. For these reasons, such investigations have almost exclusively been applied to study the penetration and the mode of action of insecticides in relatively large pest species (see O’Brien6 and Gerolt7 for reviews). The method of choice that has been extensively used consists of detecting radiolabeled compounds. In more recent studies, this method has also been successfully applied to study the insecticide residual uptake by beneficial species such as epigeal spiders8 and small parasitoids,9,10 both exposed to insecticides in laboratory conditions. However, despite the numerous analytical advantages brought by the use of radiolabeled compounds in laboratory studies, this technique is not compatible with the quantification of insecticide uptake by these insects exposed under field conditions. The development of analytical methods using chromatography techniques may allow a more systematic monitoring of beneficial insect exposure. In their study, Longley and Stark10 showed that similar amounts of diazinon could be detected in batches of five aphid parasitoids by either the detection of the radiolabeled compound or the analysis of the formulated compound in solvent extracts by GC-NPD. Therefore, by improving the extraction procedure of the analyte from the insect matrix, it may be possible to apply such techniques to the detection of the very small amounts of insecticides that may be present in individual insects. The main limitation of the solvent extraction procedure lies in the amount of the compound that is present in the injection volume (1 µL) and in the occurrence of coextracts that are not always removed by filtration and may affect the detection limits. The use of the SPME technique, which has recently been developed by (4) Stevenson, J. H.; Walters, J. H. H. Agric. Ecosyst. Environ. 1983, 10, 201215. (5) Eijsackers H. Ecotoxicology of Soils Organisms; Donker, M. H., Eijsackers, H., Heimbach, F. Eds; Setac Publication Series; Lewis Publishers: London, U.K., 1993; Chapter 1. (6) O’Brien, R. D. Insecticides: Action and Metabolism; Academic Press: New York, 1967; Chapter 16. (7) Gerolt, P. Biol. Rev 1983, 58, 233-274. (8) Jagers op Akkerhuis, G. A. J. M.; Hamers, T. M. Pestic. Sci. 1992, 36, 5968. (9) Bull, D. L.; Pryor, N. W.; King, E. G. J. Econ. Entomol. 1987, 80, 739-749. (10) Longley, M.; Stark, J. D. Bull. Environ. Contam. Toxicol. 1996, 57, 683690.
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Pawliszyn and co-workers,11,12 provides a solution to these problems. The technique exploits the affinity of analytes for a fiber coated with an adsorbent polymer to extract them from an aqueous solution. Thus, by selectively concentrating these analytes on the fiber, SPME significantly improves the sensitivity of the analysis. The theory and applications of this technique have already been described in detail.13 The method has been successfully applied to the monitoring of various insecticides in water14-16 and soil.17 SPME also proved to be suitable for extracting free amounts of a wide range of compounds in rat hepatocytes.18 In this study, we propose a method using SPME to quantify insecticide uptake in beneficial parasitoids. The method was developed using the parasitic wasp Trybliographa rapae, (Hymenoptera: Figitidae) which is a species of major importance for the control of the cabbage root fly, Delia radicum (Diptera: Anthomyiidae), a common pest in cruciferous crops.19 We envision using this beneficial insect in combination with the organophosphate chlorfenvinphos in an IPM program; therefore, it is valuable to determine the fraction of the insecticide that may be absorbed in T. rapae, as well as the fraction that may induce toxicological effects. It is also important to assess the elimination rate of the insecticide in wasps and to know whether long-term effects can be expected. The suitability of SPME to study these aspects of chlorfenvinphos toxicodynamic was investigated in topically treated wasps. The method was then used to quantify chlorfenvinphos uptake in parasitoids that had been exposed to the pesticide under field conditions. EXPERIMENTAL SECTION Apparatus and Reagents. SPME fibers were purchased from Supelco (Bellefonte, PA). Preliminary tests showed that fibers coated with 100 µm of PDMS achieved a greater extraction of chlorfenvinphos in aqueous samples than fibers coated with 85 µm of polyacrylate. The 100-µm PDMS fiber was, therefore, selected for analysis. All analyses were performed using a Carlo Erba 6000 Vega series GC ECD controlled by a ICU 600 module (Carlo Erba Instruments; Milan, Italy) and equipped with a split/splitless injector. A 30 m × 0.32 mm fused-silica DB-5 (0.25-µm film thickness) column (J&W Scientific; Folson, CA) was used. Desorption of the fiber was performed during 5 min in the injector used in the splitless mode and held at 220 °C. This temperature was maintained during the entire run. The optimized oven temperature program for chlorfenvinphos was as follows: oven kept initially at 100 °C for 1 min, temperature raised at 10 °C min-1 to 200 °C, then at 1 °C min-1 to 220 °C, this final temperature being maintained for 2 min. The detector was maintained at 300 (11) Arthur, C. L.; Pawliszyn, J. Anal. Chem. 1990, 62, 2145-2148. (12) Louch, D.; Motlagh, S.; Pawliszyn, J. Anal. Chem. 1992, 64, 1187-1189. (13) Pawliszyn, J. Solid-Phase Microextraction: Theory and Practice; John Wiley and Sons: New York, 1997; Chapter 4. (14) Eisert, R.; Levsen, K.; Wu ¨ nsch, G. J. Chromatogr. 1994, 683, 175-183. (15) Eisert, R.; Levsen, K. Fresenius J. Anal. Chem. 1995, 351, 555-562. (16) Eisert, R.; Levsen, K. J. Am. Soc. Mass. Spectrom. 1995, 6, 1119-1130. (17) Magdic, S.; Boyd-Boland, A.; Jinno, K.; Pawliszyn, J. J. Chromatogr. 1996, 736, 219-228. (18) Vaes, W. H. J.; Ramos, E. U.; Hamwijk, C.; van Holsteijn, I.; Blaauboer, B. J.; Seinen, W.; Verhaar, J. M.; Hermens, J. L. M. Chem. Res. Toxicol. 1997, 10, 1067-1072. (19) Wishart, G.; Montheit, E. Can. Entomol. 1954, 86, 143-154.
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°C. The carrier and make-up gas was nitrogen. The carrier gas flow was 1 mL min-1. Chlorfenvinphos standards were prepared using the certified compound (purity > 98%; CIL Cluzeau; Puteaux, France) in water (Milli-Q water purification system; Millipore; Pittsburgh, PA). The insecticide solutions used for the treatment of insects were prepared using the formulated compound (Birlane CE40; Cyanamid Agro; France), because formulations often contain adjuvants facilitating the penetration of chemicals across plant cuticle and, thereby, probably also across the insect cuticle. The solvents used during the insect preparation procedure for the quantification of internal doses in insects were of Normapur quality (Prolabo; Fontenay sous Bois, France). The ethyl acetate used in the definite preparation procedure was of Pestinorm quality (Prolabo). Optimization of the SPME Conditions for Chlorfenvinphos. A 1.5 µg L-1 solution of chlorfenvinphos in water was prepared. Aliquots of 7 mL were transferred to 10-mL sample vials (Thermoseparation Products; Les Ulis, France) containing 4.5 × 10 mm spin bars and stirred at a regular speed of 1000 rpm. Adsorption on PDMS fiber was performed during 1, 5, 10, 20, 40, 60, 120, or 240 min. For each adsorption time, 6 repetitions were made with a new aliquot. The amount that was thermally desorbed from the fibers during injection was calibrated using standards prepared in hexane that were analyzed by keeping the same GC conditions. The intensities of the measured signals were plotted vs sampling time and fitted using the following equation, according to Ai.20
A ) A0[1 - exp(-at)]
(1)
where A0 is the signal obtained at equilibrium and corresponds to the adsorption by the fiber of the amount n0 of chlorfenvinphos. The parameter a is a measure of how fast the equilibrium can be reached in the SPME process and is constant in systems that are agitated in a constant way.20 In our study, the time required for chlorfenvinphos to reach the equilibrium, which is assumed to be reached when 95% of n0 is adsorbed onto the fiber,13 was relatively high. A shorter sampling time was selected using the above equation. All calculations were performed using Statgraphics Plus from Manugistic Co. The linearity range of the adsorption process at the selected sampling time (45 min) was determined. Chlorfenvinphos solutions ranging from 0.05 to 50 µg L-1 were prepared. For each solution, 7 aliquots of 7 mL each were measured. Rearing and Treatment of Insects. Strains of T. rapae and D. radicum were obtained from the INRA station at Le Rheu (France). T. rapae was reared on D. radicum larvae infesting rutabaga (Brassica napus L. var. napobrassica) roots, as described in Neveu et al.21 Rearing and experiments were conducted in a climatic room [20 ( 1 °C, 60 ( 10% RH, and 16:8 h (L:D) photoperiod]. The insects used in the experiments were 0-24 h old and fed with honey prior to use. Males and females that were used were 2.94 ( 0.03 and 2.97( 0.03 mm long, respectively. Before treatment, they were isolated in plastic boxes (8 cm in diameter × 2 cm high) provided with an aeration hole located on (20) Ai, J. Anal. Chem. 1997, 69, 1230-1236. (21) Neveu, N.; Kacem, N.; Ne´non, J. P. IOBC/WPRS Bulletin 1996, 19, 173178.
the lid and covered with fine mesh. They were then immobilized by a 5-s exposure to carbon dioxide and submitted to a topical application of 0.5 µL of the required insecticide solution on the abdomen. Applications were performed using a 1-µL syringe fitted with a bevel point (Hamilton). Insects were kept in the plastic boxes and provided with water and honey until analysis. Determination of Chlorfenvinphos Uptake in Insects. Because of the possible complexity in the distribution of chlorfenvinphos between the different components of the insect matrix, different insect preparation procedures were investigated prior to the fiber extraction. Insects (all females) used in this experiment were treated with a nonlethal dose of chlorfenvinphos (30 ng/insect)22 and kept in a box for 1 h prior to analysis. This delay was observed to allow insecticide penetration through the cuticle; this time was short enough to avoid possible losses of insecticide through elimination or metabolism.6 Insects were then killed by placing them for a few minutes at - 18 °C. They were then transferred individually to Eppendorf tubes and rinsed in 200 µL ethyl acetate to determine the fraction of chlorfenvinphos remaining on the cuticle, and, thus, the penetration rate. Rinsing volumes were transferred to sampling vials, dried under a nitrogen flow, and recovered in 7 mL of water for microextraction and analysis. To quantify the free concentration of chlorfenvinphos in their body, which also corresponds to the toxicological concentration, insects were crushed manually in 200 µL of water using potter Kontes homogenizers that tightly fitted the Eppendorf tubes. The mixtures were then left for 1 h at 20 °C to allow a reproducible solubilization of the insecticide. Tubes were then centrifuged for 10 min at 800 rpm to separate the cuticle from the insect matrix. Cuticles were rinsed 3 times with 200 µL of water. All of the supernatants were transferred in sampling vials and adjusted to 7 mL with water for the SPME step. The influence of the insect matrix on the adsorption process was investigated by spiking untreated extracts with 30 ng of chlorfenvinphos in water. The total uptake of chlorfenvinphos in insects, which is directly related to the exposure, was also determined. This experiment was performed by the replacement of water by an organic solvent during the crushing of insects to solubilize a maximum amount of the insecticide. Ethyl acetate, acetone, acetonitrile, and dichloromethane, which are commonly used to extract chlorfenvinphos from biological matrixes,10,23,24 were tested as possible crushing solvents. As for water-prepared extracts, the influence of the insect matrix on the adsorption process was investigated by spiking untreated extracts with 30 ng of chlorfenvinphos in water. Finally, the influence of sonication on chlorfenvinphos recovery was also tested on water and ethyl acetate extracts. Extracts were sonicated for 3 min using a sonication cell (Vibra cell; Bioblock; Illkirch, France). As for water-prepared extracts, the influence of the insect matrix on the adsorption process was investigated by spiking untreated extracts with 30 ng of chlorfenvinphos in water. Each procedure was repeated 10 times in order to study the reproduc(22) Alix, A.; Cortesero, A. M.; Ne´non, J. P.; Anger, J. P. Environ. Toxicol. Chem., submitted. (23) Ikeda, T.; Tsuda, S.; Shirasu, Y. Fundam. Appl. Toxicol. 1991, 17, 361367. (24) Serrano, R.; Lo´pez, F. J.; Herna´ndez, F.; Pen ˜a, J. B. Bull. Environ. Contam. Toxicol. 1997, 59, 968-975.
Figure 1. Adsorption profile of chlorfenvinphos in water.
ibility of the recovery rate and the possible effects of the preparation method on the lifetime of the SPME fiber. A new fiber was used for each new procedure (10 repetitions). Elimination Kinetics of Chlorfenvinphos in T. rapae. Insects of both sexes were submitted to the application of 20 ng of chlorfenvinphos and killed 2, 24, 60, 120, 240 and 360 h after treatment. The distribution of the insecticide between the body fat and the haemolymph may evolve during the insect’s lifetime, so that the body-fat fraction may be at least partially involved in the amounts that are eliminated. To also take into account this fraction, insect extracts were prepared in ethyl acetate and sonicated. Each measure was repeated 7 times for both sexes. Chlorfenvinphos uptake by insects exposed under field conditions. D. radicum pupae were collected in a cabbage field (St Me´loir des Ondes, France) in which plants had been treated with chlorfenvinphos (Birlane CE40) by root dipping at the time of planting. The pupae were collected 21 days after planting, which corresponds to the first generation of insects that developed from the treated plants. Parasitoids emerging from these pupae were prepared as described above, using either ethyl acetate or water as the crushing solvent in order to determine the doses and their free fraction that could be absorbed by insects during their development in field conditions. In these samples, the presence of chlorfenvinphos was confirmed by mass spectrometry. RESULTS AND DISCUSSION Optimization of the Extraction Conditions for Chlorfenvinphos. The adsorption profile for chlorfenvinphos in an aqueous solution is shown in Figure 1. The line corresponds to the adsorption model given by eq 1, which gives an accurate description of the adsorption process (A ) 358927[1 exp(- 0.021134t)], r2 ) 0.9967). According to this model, a 142min sampling time would be necessary to reach the adsorption equilibrium. Nonetheless, it has already been demonstrated that a quantitative analysis can be performed even in nonequilibrium situations, provided that adsorption time and agitation conditions are carefully controlled and kept constant during the adsorption process.11,20 In such cases, the sampling time should be selected so that a deviation of less than 5% in the adsorbed analyte is observed for a 1-min error.13,25 In our study, a 45-min adsorption time was selected in order to achieve a signal/noise ratio of 10 for amounts of ∼0.5 ng in the vial and, thus, to provide a quantification sensitivity that is suitable to insect uptake. At this (25) Vaes, W. H. J.; Ramos, E. U.; Verhaar, H. J. M.; Seinen, W.; Hermens, J. L. M. Anal. Chem. 1996, 68, 4463-4467.
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Table 1. Influence of Solvent and Sonication on the Recovery Rate of Chlorfenvinphos in Insect Extractse solvent
sonication
amt (ng) detected in extracts, % RSD
recovery, %
water hexane acetone acetonitrile dichloromethane ethyl acetate water ethyl acetate waterf ethyl acetatef
no no no no no no yes yes yes yes
15.63 (21.54)a 14.76 (30.84)a 21.90 (23.39)ab 16.98 (22.18)ad 24.40 (13.65)bc 22.98 (15.38)bcd 19.84 (11.01)ac 25.32 (6.37)b 28.64 (4.11) 29.00 (3.46)
52 49 73 57 81 77 66 84 95 97
e Values followed by the same letter are not significantly different at the 0.001 level (t-tests, 18 df) f Insect matrix spiked with 30 ng chlorfenvinphos.
time, the deviation of the quantity adsorbed was only 1.30% for an uncertainty of 1 min, and 61.36% of n0 were adsorbed. This amount corresponds to 2.02% of the amount of chlorfenvinphos that is contained in the vial. Using this 45-min sampling time, the adsorption of chlorfenvinphos was linear from 0.07 to 15 ng mL-1 (r2 ) 0.997, F ) 12877.5, P < 0.001, df ) 41). The detection limit was 0.02 ng mL-1, using a signal/noise ratio of 3. Determination of Chlorfenvinphos Uptake in Insects. About 52% of the insecticide applied to insects was retrieved from the matrixes prepared in water, with a standard deviation of 21.54% (Table 1). The recovery was raised to 66% by an additional sonication of the matrix during the crushing step, but this gain was not significant. The complete lysis of insect tissues and cells that occurs may enhance the homogeneity of the matrix, thus facilitating the adsorption of the insecticide onto the fiber. The lysis of cells may also lead to a release into water of the fraction of the insecticide that is retained in the cellular compartment. These phenomena may explain why the variability in these samples was halved, although this decrease was not significant (df ) 18, F ) 2.42, P ) 0.10). However, sonication may not significantly displace the equilibrium between bound and free forms of the insecticide. Thus, the fraction of the insecticide that was extracted from these samples may well correspond to the free dissolved chlorfenvinphos only, because lower amounts were adsorbed onto the fiber from these mixtures than from those that had been prepared in ethyl acetate and sonicated (Table 1). Moreover, as suggested by Vaes et al.,25 the equilibrium of an analyte between water and a matrix is not displaced by the extraction of small amounts of the analyte from the water volume. In our study, only 2.02% of the freely dissolved chlorfenvinphos were extracted by the SPME fiber; therefore, it can be assumed that the amount of chlorfenvinphos that was extracted by fibers in these samples is the fraction of the insecticide that was freely dissolved in the insect body fluids. The recovery of chlorfenvinphos in the spiked insect matrixes was very high (Table 1), showing that the biological components present in the vial did not interfere with the adsorption of the insecticide onto the PDMS coating. This proves that a quantification of the freely dissolved chlorfenvinphos in insect bodies can be performed using SPME. 3110 Analytical Chemistry, Vol. 73, No. 13, July 1, 2001
The quantification of the total uptake of chlorfenvinphos by parasitoids provided variable results, depending on the crushing solvent (Table 1). The amounts of chlorfenvinphos solubilized by acetone, acetonitrile, or hexane were not reproducible and, thereby, were not found to be different from the amounts solubilized in water. Ethyl acetate and dichloromethane provided the higher recovery rates, because 77 and 81%, respectively, of the applied doses were detected. However, the fibers used to extract chlorfenvinphos from matrixes prepared in dichloromethane showed a rapid aging, visible as losses of coating, and the use of this solvent was subsequently abandoned. Ethyl acetate seemed to offer the best compromise between extraction and reproducibility. The increase in reproducibility that was provided by an additional sonication step was significant (df ) 18, F ) 4.83, P < 0.05), suggesting an improved homogenization of the matrix. In these conditions, the recovery of chlorfenvinphos was 84% of the applied dose, and the precision was 6.37%. As for water-prepared spiked samples, the extraction of chlorfenvinphos from ethyl acetate-spiked samples provided an almost complete recovery, showing that the insect matrix modified by the action of ethyl acetate did not interfere with the adsorption process of the insecticide on the SPME fiber. Therefore, the results achieved by this method can be considered as the total amount of chlorfenvinphos that could be solubilized from the insect matrix by solvent. The gain of recovery provided by the use of this solvent may correspond to the solubilization of the amount of insecticide immobilized in the insect body.26 The hydrophobic properties of the chemical, as suggested by its octanol-water coefficient (log Kow ) 3.85) support this hypothesis. The analysis of the external rinses of the wasps showed that the residues remaining on the insect cuticle were systematically lower than the detection limit (0.02 ng mL-1). This result suggests a rapid and complete penetration of chlorfenvinphos through the insect integument, because ∼99.5% of the insecticide was absorbed into the insects within an hour. The affinity of the insecticide for organic matrixes (log Kow) may in part explain such diffusion properties. Baur et al.27 found that the high diffusion capacity of chlorfenvinphos through plant cuticles was related to the plasticizing effect of the chemical on the cuticular waxes. Despite the fact that the parallel between plant and insect cuticles is difficult to draw, this result outlines the strong interaction capacity of this chemical. This high penetration rate of the insecticide through parasitoids’ integument suggests that the amount of the insecticide that was not retrieved in ethyl acetate extracts may in part be retained in the insect cuticle. In previous work, Gerolt7 showed that a significant fraction of radiolabeled organophosphorus compounds could be strongly immobilized into the integument so that they were not extractable in solvents. The author also noted that this immobilization might not be definite, because chemicals were able to slowly diffuse into the cuticular layers to finally pass through the epidermis into the haemolymph. Recent modeling studies on the diffusion kinetics of insecticides through insect integument resulted in similar conclusions.28 Thus, although the cuticular fraction may not be easily bioavailable, this must be taken into account when attempting to determine the total uptake of an (26) Seydel, J. K.; Schaper, K. J. Pharmacol. Ther. 1982, 15, 131-182. (27) Baur, P.; Grayson, B. T.; Scho ¨nherr, J. Pestic. Sci. 1996, 47, 171-180. (28) Salgado, V. L. Pestic. Sci. 1995, 44, 59-67.
Table 2. Internal Dose of Chlorfenvinphos in Adult Trybliographa rapae Emerging from Host (Delia radicum) Pupae Collected in a Treated Cabbage Field amt (ng) chlorfenvinphos detected, % RSD
water ethyl acetate
Figure 2. Elimination kinetic of chlorfenvinphos in adult parasitoids. Bars represent the standard errors.
insecticide. Then the amount solubilized by the solvent corresponds more precisely to the fraction partitioned between fat tissues and body fluids, and represents the internal dose at a defined time. In the case of T. rapae exposed to chlorfenvinphos, this internal dose may be estimated at about 84% of the dose contacted, among which 78% remains freely dissolved. Therefore, up to 66% of the dose contacted may pose a toxicological risk for the parasitoid within the first hour after exposure. Using SPME, this freely dissolved fraction may be quantified up to 0.5 ng/insect, which corresponds to an uptake of 0.75ng. Such a quantification would not have been possible using a conventional solvent extraction process. The sensitivity of the present method may still be improved by the use of smaller sample volumes, as suggested by Louch et al..12 Elimination Kinetics of Chlorfenvinphos in T. rapae. The internal doses of chlorfenvinphos detected in parasitoids over time are depicted in Figure 2. The elimination of the insecticide was very slow within the first days after treatment and became significant after 48 h in females (t ) 2.039, P < 0.05, df ) 12) and 120 h (5 days) in males (t ) 2.097, P < 0.05, df ) 12). No differences were found in the internal doses retrieved between males and females, which suggests that the same mechanism(s) are (were) involved in the elimination of the insecticide. Metabolism and excretion are the two main mechanisms by which toxicants may be eliminated, assuming that storage only immobilizes them for a defined time. Because of the hydrophobic properties of this chemical, it is likely that elimination through excretion may have required a prior transformation into a more polar compound. Because the turnover of many degrading enzymatic systems is very low,6 the elimination process may take a relatively long time, which may explain the above results. According to the low elimination rate of chlorfenvinphos in this parasitoid, it is likely that additional contact with the insecticide would lead to accumulation in the insect body. These results are in agreement with the bioconcentration properties of chlorfenvinphos already described in mussels.23 These considerations may be of great importance when assessing the risks posed by an insecticide for nontarget insects, because they are continuously exposed in treated fields.
males
females
8.21 (79.51) (n ) 8) 9.30 (106.08) (n ) 13)
5.93 (77.95) (n ) 9) 7.41 (88.02) (n ) 11)
Chlorfenvinphos Uptake by Insects Exposed under Field Conditions. Significant amounts of chlorfenvinphos were detected in parasitoids collected from a treated field (Table 2). Because the development of these parasitoids takes place inside the body of their hosts, these amounts must have been absorbed via the host. The great variability of the doses that were absorbed (from 0.36 to 19.50 ng in females, and from 0.45 to 26.06 ng in males) may be in part related to the variability of the doses that the hosts may absorb while feeding on the treated plants. Because of the important variability of the uptake between insects, no difference was found between the amounts that were detected using water and those that were retrieved using ethyl acetate during the preparation. However, the freely dissolved fraction of chlorfenvinphos retrieved in the insects may be conformed to toxicological data. The median lethal doses of chlorfenvinphos we determined in topically treated wasps, 24 h after treatment, were 387 ( 26 ng/insect and 690 ( 54 ng/insect in males and females, respectively.22 From our results, it can be assumed that two-thirds of these doses (255 and 455 ng) may have been responsible for the lethal effects. Thus, the freely dissolved amounts of chlorfenvinphos in insects emerging in a treated field are far lower than their median sensitivity to lethal effects. The risk related to chlorfenvinphos bioconcentration properties in such insects, while contacting other treated plants to find their hosts, now needs to be investigated. CONCLUSION The use of SPME as a sampling technique enables the quantification of the very low amounts of chlorfenvinphos that may be contained in small species of beneficial parasitoids (from 2.19 to 3.70 mm long for this species). The sensitivity of the method also permits quantification of the amounts of chemical that are really involved in toxicological effects, including sublethal effects. This method, finally, proved to be suitable for determining both free and total internal amounts of the insecticide in field-exposed parasitoids, thus offering new possibilities for monitoring studies. ACKNOWLEDGMENT This work was supported by the Conseil Re´gional de Bretagne. We thank A.M. Cortesero, D. Poinsot, and J. O. Stapel for their helpful comments. Received for review November 8, 2000. Accepted April 3, 2001. AC0013148
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